One of the most common problems in chemical and biochemical research is to separate components from each other, for example larger molecules from smaller ones, macromolecules in cell extracts and the like. Methods for separating components of a mixture exploit differences in properties of the components, such as in size, electrical charge, solubility in different solvents, affinity and hydrophobic interactions.
Exclusion chromatography refers to a technique in which the separation is based mainly on exclusion effects, such as differences in molecular size and/or shape and/or in charge, and it includes size-exclusion chromatography (SEC).
Size-exclusion chromatography (SEC), also called gel-filtration chromatography (GFC) or gel-permeation chromatography (GPC) uses porous particles to separate molecules of different sizes when the stationary phase is a swollen gel. It is usually applied to large molecules or macromolecular complexes such as proteins and industrial polymers. Typically, when an aqueous solution is used to transport the sample through the column, the technique is known as GFC, versus the name GPC when an organic solvent is used as a mobile phase. SEC is also used for polymer characterization.
The main application of gel-filtration chromatography is the fractionation of proteins and other water-soluble polymers, while gel permeation chromatography is used to analyze the molecular weight distribution of organic-soluble polymers.
SEC is used for the purification and analysis of synthetic and biological polymers, such as proteins, polysaccharides and nucleic acids. Biologists and biochemists typically use a gel medium, usually polyacrylamide or agarose and filter under low pressure. Polymer chemists use either a silica or cross-linked polystyrene medium under a higher pressure. These media are known as the stationary phase.
SEC can be used to determine molecular weights and molecular weight distributions of polymers and to give polymer chemists information on the polydispersity of the sample. The preparative SEC can be used for polymer fractionation on an analytical scale.
Molecules that are smaller than the pore size of the stationary phase particles can enter the particles and therefore have a longer path and longer transit time than larger molecules that cannot enter the particles. SEC is a chromatographic technique that employs specialized columns to separate natural and synthetic polymers, biopolymers, proteins and nanoparticles on the basis of size. A column is filled with semi-solid beads of a polymeric gel that will admit ions and small molecules into their interior but not large ones. When a mixture of molecules and ions dissolved in a solvent is applied to the top of the column, the smaller molecules (and ions) are distributed through a larger volume of solvent than is available to the large molecules. Consequently, the large molecules move more rapidly through the column, and in this way the mixture can be separated (fractionated) into its components. The porosity of the gel can be adjusted to exclude all molecules above a certain size.
Molecules larger than the pore size cannot enter the pores and elute together as the first peak in the chromatogram. This condition is called total exclusion. Molecules that can enter the pores will have an average residence time in the particles that depends on the molecules size and shape. Different molecules therefore have different total transit times through the column. This portion of a chromatogram is called the selective permeation region. Molecules that are smaller than the pore size can enter all pores, and have the longest residence time on the column and elute together as the last peak in the chromatogram. This last peak in the chromatogram determines the total permeation limit.
One requirement for SEC is that the analyte does not interact with the surface of the stationary phases. Differences in elution time are based solely on the volume the analyte “sees”. Thus, a small molecule that can penetrate every corner of the pore system of the stationary phase “sees” the entire pore volume and the interparticle volume, and will elute late (when the pore- and interparticle volume has passed through the column ˜80% of the column volume). On the other extreme, a very large molecule that cannot penetrate the pore system “sees” only the interparticle volume (˜35% of the column volume) and will elute earlier when this volume of mobile phase has passed through the column. The underlying principle of SEC is that particles of different sizes will elute (filter) through a stationary phase at different rates. This results in the separation of a solution of particles based on size. Provided that all the particles are loaded simultaneously or near-simultaneously, particles of the same size should elute together.
Each size exclusion column has a range of molecular weights that can be separated. The exclusion limit defines the molecular weight at the upper end of this range and is where molecules are too large to be trapped in the stationary phase. The permeation limit defines the molecular weight at the lower end of the range of separation and is where molecules of a small enough size can penetrate into the pores of the stationary phase completely and all molecules below this molecular mass are so small that they elute as a single band.
Also, typically the stationary-phase particles are not ideally defined; both particles and pores may vary in size. The stationary phase may also interact in undesirable ways with a particle and influence retention times, though great care is usually taken to use stationary phases that are inert and minimize this issue.
Electrophoresis is a method which separates macromolecules by their charge and size. In gel electrophoresis an electric field is used to “pull” or “push” molecules through the gel, depending on their electrical charges. “Electrophoresis” refers to the electromotive force (EMF) that is used to move the molecules through the gel matrix. Using an electric field, molecules (such as DNA) can be made to move through a gel. The molecules being sorted are dispensed into a well in the gel material. The gel is placed in an electrophoresis chamber, which is then connected to a power source. By applying an electric field, the molecules will move through the matrix at different rates, determined largely by their mass when the charge to mass ratio (Z) of all species is uniform, toward the anode if negatively charged or toward the cathode if positively charged. The different sized molecules form distinct bands on the gel.
Gel electrophoresis refers to using a gel as an anticonvective medium and/or sieving medium, which can also be called stationary phase, during electrophoresis. The term “gel” in this instance refers to the matrix used to contain then separate the target molecules. In most cases, the gel is a cross-linked polymer whose composition and porosity is chosen based on the specific weight and composition of the target to be analyzed. When separating proteins or small nucleic acids the gel is usually composed of different concentrations of acrylamide and a cross-linker, producing different sized mesh networks of polyacrylamide. When separating larger nucleic acids, the preferred matrix is purified agarose. In both cases, the gel forms a solid, yet porous matrix. Acrylamide, in contrast to polyacrylamide, is a neurotoxin and must be handled using appropriate safety precautions to avoid poisoning.
After the electrophoresis is complete, the molecules in the gel can be stained to make them visible. For example, ethidium bromide, silver, or Coomassie brilliant blue dye may be used for this process. Other methods may also be used to visualize the separation of the mixture's components on the gel. If the analyte molecules fluoresce under ultraviolet light, a photograph can be taken of the gel under ultraviolet lighting conditions, often using a Gel Doc. If the molecules to be separated contain radioactivity added for visibility, an autoradiogram can be recorded of the gel.
If several samples have been loaded into adjacent wells in the gel, they will run parallel in individual lanes. Depending on the number of different molecules, each lane shows separation of the components from the original mixture as one or more distinct bands, one band per component. Incomplete separation of the components can lead to overlapping bands, or to indistinguishable smears representing multiple unresolved components.
Bands in different lanes that end up at the same distance from the top contain molecules that passed through the gel with the same speed, which usually means they are approximately the same size. There are molecular weight size markers available that contain a mixture of molecules of known sizes. If such a marker was run on one lane in the gel parallel to the unknown samples, the bands observed can be compared to those of the unknown in order to determine their size. The distance a band travels is approximately inversely proportional to the logarithm of the size of the molecule.
Gel electrophoresis is most commonly used for separation of biological macromolecules such as deoxyribonucleic acid (DNA), ribonucleic acid (RNA), or protein; however, gel electrophoresis can be used for separation of nanoparticles as well. Gels suppress the thermal convection caused by application of the electric field, and can also act as a sieving medium, retarding the passage of molecules; gels can also simply serve to maintain the finished separation, so that a post electrophoresis stain can be applied. DNA Gel electrophoresis is usually performed for analytical purposes, often after amplification of DNA via PCR, but may be used as a preparative technique prior to use of other methods such as mass spectrometry, RFLP, PCR, cloning, DNA sequencing, or Southern blotting for further characterization.
Gel electrophoresis is typically used in forensic, molecular biology, genetics, microbiology and biochemistry applications. The results can be analyzed quantitatively by visualizing the gel with UV light and a gel imaging device. The image is recorded with a computer operated camera, and the intensity of the band or spot of interest is measured and compared against standard or markers loaded on the same gel. The measurement and analysis are mostly done with specialized software.
Depending on the type of analysis being performed, other techniques are often implemented in conjunction with the results of gel electrophoresis, providing a wide range of field-specific applications.
SDS-PAGE, sodium dodecyl sulfate polyacrylamide gel electrophoresis, is a technique widely used particularly in forensic, molecular biology, genetics, and biochemistry to separate components according to their electrophoretic mobility (a function of length of polypeptide chain or molecular weight). SDS gel electrophoresis of samples that have identical charge per unit mass due to binding of SDS, results in fractionation by size.
Two-dimensional gel electrophoresis, abbreviated as 2-DE or 2-D electrophoresis, is a form of gel electrophoresis commonly used to analyze proteins. Mixtures of proteins are separated by two properties in two dimensions on 2D gels.
Temperature Gradient Gel Electrophoresis (TGGE) and Denaturing Gradient Gel Electrophoresis (DGGE) are forms of electrophoresis which use either a temperature or chemical gradient to denature the sample as it moves across an acrylamide gel. TGGE and DGGE can be applied to nucleic acids such as DNA and RNA, and (less commonly) proteins. TGGE relies on temperature dependent changes in structure to separate nucleic acids. DGGE was the original technique, and TGGE a refinement of it.
Agar is a linear and non-ionic polysaccharide consisting of D-galactose and 3,6-anhydro-L-galactose and it is produced from seaweeds. Agarose based electrophoresis gels are typically used in molecular biology laboratories, in methods comprising the separation of various sized DNA fragments, which are thereafter cut out from the gel, isolated and subsequently used for various purposes. The use of agarose requires warming of the agarose suspension before casting of the gel. However, warming destroys the intercalating dye which is added to the gel for DNA detection. Further, for isolating a desired DNA fragment from the gel it has to be cut mechanically out from the gel under strong UV light and in the presence of the dye. Further, in order to separate the DNA fragments from the gel for further use the gel must be removed.
Other gels used earlier for electrophoresis include starch gels.
Polyacrylamide gels are commonly used for electrophoresis particularly in the separation of nucleic acids. In general, stock solutions containing acrylamide monomer, a crosslinker such as bisacrylamide, gel buffers, and modifying agents such as sodium dodecyl sulphate (“SDS”) are prepared. To manufacture a gel, the stock solutions are mixed with water in proportions according to the final desired concentrations of the various constituents and the polymerization reaction is effected.
The resolving power of the gel is determined by the content of agarose in the gel in the case of agarose gels. In contrast, the resolving power of polyacrylamide gels is adjusted by the mixing ratio of acrylamide to bisacrylamide.
US 2010/0236932 publication proposes composite gels comprising acrylamide or N-modified acrylamide, agarose, buffers and photo-initiators with or without cross-linkers.
The detection of nucleic acids in nucleic acid analytics with the aid of gel electrophoresis is based on the fact that fluorescent DNA binding dyes permanently bind non-covalently to nucleic acids and, in their bound form, enable nucleic acids to be located in the gel matrix after excitation with light of a suitable wavelength.
Ethidium bromide is typically used as the intercalating dye, however, it is neurotoxic and carcinogenic. Due to the toxicity, alternative DNA binding dyes such as for example SYBR Green or SYBR Gold (Molecular Probes, Inc.) are also used, the binding properties of which are not or not exclusively based on the principle of DNA intercalation.
The dyes used for electrophoresis are either admixed with the gel preparation before polymerization in the case of ethidium bromide, or the gel is stained after completion of the gel electrophoresis with the aid of an aqueous dye solution containing ethidium bromide or another dye. Furthermore, SYBR Green I can be added to the sample containing the nucleic acid before loading the gel.
Ion exchange chromatography is a popular method for the purification of proteins and other charged molecules. In cation exchange chromatography positively charged molecules are attracted to a negatively charged solid support (stationary phase). Conversely, in anion exchange chromatography, negatively charged molecules are attracted to a positively charged solid support.
To optimize binding of all charged molecules, the mobile phase is generally a low to medium conductivity solution. The adsorption of the molecules to the solid support is driven by the ionic interaction between the oppositely charged ionic groups in the sample molecule and in the functional group on the support. The strength of the interaction is determined by the number and location of the charges on the molecule and on the functional group. By increasing the salt concentration the molecules with the weakest ionic interactions start to elute from the column first. Molecules that have a stronger ionic interaction require a higher salt concentration and elute later in the gradient. The binding capacities of ion exchange resins are generally quite high. This is of major importance in process scale chromatography, but is not critical for analytical scale separations.
As a rule, the pH of the mobile phase buffer must be between the pI (isoelectric point) or pKa (acid dissociation constant) of the charged molecule and the pKa of the charged group on the solid support. For example, in cation exchange chromatography, using a functional group on the solid support with a pKa of 1.2, a sample molecule with a pI of 8.2 may be run in a mobile phase buffer of pH 6.0. In anion exchange chromatography a molecule with a pI of 6.8 may be run in a mobile phase buffer at pH 8.0 when the pKa of the solid support is 10.3.
For example, a protein sample is injected onto the column under conditions where it will be strongly retained. A gradient of linearly increasing salt concentration is then applied to elute the sample components from the column. An alternative to using a linear gradient is to use a step gradient. This requires less complicated equipment and can be very effective to elute different fractions if the appropriate concentrations of salt are known, usually from linear gradient experiments.
Also changes in pH are used to affect separation. In cation exchange chromatography, raising the pH of the mobile phase buffer will cause the molecule to become less protonated and hence less positively charged. The result is that the protein no longer can form ionic interaction with the negatively charged solid support, which ultimately results in the molecule to elute from the column. In anion exchange chromatography, lowering the pH of the mobile phase buffer will cause the molecule to become more protonated and hence more positively charged. The result is that the protein no longer can form ionic interaction with the positively charged solid support which causes the molecule to elute from the column.
Affinity chromatography is a method of separating biochemical mixtures and it is based on a highly specific interaction such as that between antigen and antibody, enzyme and substrate, or receptor and ligand. The immobile (stationary) phase is typically a gel matrix, often of agarose. Usually the starting point is an undefined heterogeneous group of molecules in solution, such as a cell lysate, growth medium or blood serum. The molecule of interest will have a well known and defined property which can be exploited during the affinity purification process. The process itself can be thought of as an entrapment, with the target molecule becoming trapped on a solid or stationary phase or medium. The other molecules in solution will not become trapped as they do not possess this property. The solid medium can then be removed from the mixture, washed and the target molecule released from the entrapment in a process known as elution. Possibly the most common use of affinity chromotography is for the purification of recombinant proteins. Binding to the solid phase may be achieved by column chromatography whereby the solid medium is packed onto a column, the initial mixture run through the column to allow setting, a wash buffer run through the column and the elution buffer subsequently applied to the column and collected. Alternatively binding may be achieved using a batch treatment, by adding the initial mixture to the solid phase in a vessel, mixing, separating the solid phase (for example), removing the liquid phase, washing, re-centrifuging, adding the elution buffer, re-centrifuging and removing the eluate.
Sometimes a hybrid method is employed, the binding is done by the batch method, then the solid phase with the target molecule bound is packed onto a column and washing and elution are done on the column.
A third method, expanded bed adsorption, which combines the advantages of the two methods mentioned above, has also been developed. The solid phase particles are placed in a column where liquid phase is pumped in from the bottom and exits at the top. The gravity of the particles ensure that the solid phase does not exit the column with the liquid phase.
Affinity columns can be eluted by changing the ionic strength through a gradient. Salt concentrations, pH, pI, charge and ionic strength can all be used to separate or form the gradient to separate.
Affinity chromatography can be used in a number of applications, including nucleic acid purification, protein purification from cell free extracts, and purification from blood.
Another use for the procedure is the affinity purification of antibodies from blood serum. If serum is known to contain antibodies against a specific antigen (for example if the serum comes from an organism immunized against the antigen concerned) then it can be used for the affinity purification of that antigen. This is also known as immunoaffinity chromatography.
Immobilized metal ion affinity chromatography (IMAC) is based on the specific coordinate covalent bond of amino acids, particularly histidine, to metals. This technique works by allowing proteins with an affinity for metal ions to be retained in a column containing immobilized metal ions, such as cobalt, nickel, copper for the purification of histidine containing proteins or peptides, iron,zinc or gallium for the purification of phosphorylated proteins or peptides. Many naturally occurring proteins do not have an affinity for metal ions, therefore recombinant DNA technology cab be used to introduce such a protein tag into the relevant gene. Methods used to elute the protein of interest include changing the pH, or adding a competitive molecule, such as imidazole.
Based on the above it can be seen that stationary phases are necessary in several separation methods. In the methods a stationary phase provides means for separating components in an efficient way from a mixture which is conducted through the stationary phase, yet not interacting with the components in an undesired way. There is an evident need for improved stationary phases useful in separation methods, particularly in methods based on electrophoresis or chromatography.